Our goal is to determine the effects of the cell cycle, oncogenic activity, and topo IIα protein levels on the outcome of treatment with the anti-topo II drug etoposide. It was first necessary to determine exactly how topo IIα protein levels are affected by the cell cycle and oncogene action. This required a detailed cell cycle analysis of topo IIα protein levels. Our approach was to determine the cell cycle positions of individual cells in an asynchronous culture. This approach not only allowed analysis of cells in all cell cycle phases simultaneously but also eliminated any potential complication associated with methods of cell cycle synchronization. The cell cycle phase was determined in two separate ways. In the first, cells were fixed and DNA was stained with DAPI. Previous studies have conclusively demonstrated that the fluorescence intensity of the cell stained in this way is proportional to DNA content. Moreover, the analysis of monolayer cells was found to be extremely accurate due to the high-resolution optics involved, the ability to analyze fluorescence specifically within the nuclear region, and the low level of cytoplasmic interference within the nuclear region in flattened monolayer cells (20
). In the second technique, cell cycle position was assessed by monitoring cells in time-lapse for 24 h and then determining the age of each cell (or the time since passing through mitosis) at the time of fixation (21
To determine the level of topo IIα protein in each cell, the cells were stained with an antibody specific to topo IIα. It was first necessary to demonstrate that this staining procedure resulted in fluorescence levels proportional to those of topo IIα protein within the cell. To accomplish this, topo IIα protein levels and fluorescence were compared in cells synchronized by serum deprivation and restimulation. At various times following serum addition to quiescent NIH 3T3 cultures, lysates were collected for Western analysis, while cells in parallel cultures were fixed and stained for topo IIα and DNA. The amount of topo IIα protein at each time point was determined by quantitating the intensity of the Western band. This was then compared to fluorescence intensity by measuring the fluorescence of 200 to 400 cells in a parallel culture and determining the average fluorescence (21
). The results of a typical analysis (Fig. ) indicate a close correlation between the average fluorescence intensity of the topo IIα stain and the absolute amount of topo IIα protein within the culture. In separate analyses, it was found that only full-length topo IIα was present in isolated nuclei, thus reducing the possibility that the results obtained reflected partially degraded protein (data not shown). It is, therefore, possible to measure fluorescence intensity and determine the relative amounts of topo IIα in individual cells.
Increase of topo IIα protein through the cell cycle.
The levels of topo IIα through the cell cycle of actively cycling cells were next determined by fixing and staining an asynchronous culture of NIH 3T3 cells for topo IIα and DNA as described above. Separate images of each fluorochrome were collected from the same cells. The nuclear region of each cell was identified from the DNA stain, after which the total amounts of DNA- and topo IIα-associated fluorescence for each nucleus were determined. When the DNA fluorescence levels for individual cells are plotted versus topo IIα-associated fluorescence, it can be seen that the amounts of topo IIα continuously increased as the cells progress from G1 to G2 phase (Fig. A). While the amounts of topo IIα protein increase during passage through the cell cycle, it was somewhat surprising, based on previously reported data, that the increase was not larger than the one observed. Most cell constituents passively increase in amount during the cell cycle due merely to the increase in cell size and mass. It is critical to determine if this might be the explanation for the slight increase in topo IIα protein observed above.
FIG. 2 Topo IIα expression through the cell cycle. (A) An asynchronous culture of NIH 3T3 cells was fixed and stained for DNA and topo IIα. The levels of each fluorochrome were determined by image analysis, and the DNA levels for each cell were (more ...)
To test this possibility, the total amount of topo IIα fluorescence in each cell was divided by the DNA-associated fluorescence for that cell. This normalized topo IIα value was then plotted versus that for DNA. When the amount of topo IIα protein relative to DNA content was displayed in this way, it was clear that the increase in topo IIα protein through the cell cycle was roughly equal to the increase in DNA content within the cell (Fig. B). The increase in topo IIα levels was, therefore, reflective only of the overall increase in cell size as cells progress through the cell cycle. (A very few cells with high topo IIα levels during G1 and G2 phases can largely be accounted for by the presence in these populations of rounded cells in mitosis, which give artificially high fluorescence values [Fig. ].) This type of analysis was extended to a number of other cell types, including tumor cells, with identical results.
To determine if there might be a slight cell cycle-related alteration in topo IIα protein, results from nine separate experiments were compared. For this comparison, cells within each experiment were grouped according to DNA content and the average topo IIα/DNA ratio for each of these groups was determined. Data from all nine separate experiments were then compared to determine the mean topo IIα/DNA ratio (± SE) for each DNA range. When this mean value was plotted versus the DNA level, the possibility that a slight reduction in the topo IIα/DNA ratio might exist during S phase became apparent (Fig. A). While the difference was only approximately 10%, it might indicate a larger difference in the size of a particular cellular pool of topo IIα.
Time-lapse analysis of topo IIα levels.
To confirm the above results, topo IIα expression through the cell cycle was analyzed using time-lapse to determine cell cycle position. Cells of an asynchronous culture were monitored for 25 h prior to fixation and staining for topo IIα. The level of topo IIα-associated fluorescence was determined for each cell as described above. Individual cells were then monitored in the time-lapse movie to determine their ages, or how long prior to fixation they had passed through mitosis (29
). In previous studies we have shown that most NIH 3T3 cells less than 5 h old are in G1
phase, that cells 5 to 12 h old are in S phase, that cells older than 12 h are most likely in G2
phase, and that the average generation time was 16 to 17 h (20
). As an indication of the cell cycle characteristics of these cultures, the ages of individual cells in a typical analysis are plotted versus their DNA contents (Fig. A). Although most cells in the culture behave as described above, a small number display slower transit through the cell cycle. The topo IIα-associated fluorescence also increased with age (Fig. B). To relate topo IIα levels to DNA levels as these cells progressed through the cell cycle, cells were grouped according to age and the average topo IIα fluorescence levels for each group were determined, as was the average DNA fluorescence level. When the average topo IIα and the average DNA levels were plotted versus age, it was clear that topo IIα levels and DNA content increased together through the cell cycle in this time-lapse analysis (Fig. C). Based on all the above data, therefore, we conclude that the topo IIα content of cycling cells increases roughly in proportion to the increase in DNA content through the cell cycle in all cell types analyzed.
Ras injection and topo IIα levels.
We next analyzed the effects of constitutive oncogenic signaling on topo IIα levels. Oncogenic Ras (L61; 1 mg/ml) was microinjected into cells of an asynchronous culture, and the topo IIα and DNA levels were determined 24 h later by fluorescence photography and image analysis. No increase in the topo IIα levels was observed following Ras injection during any cell cycle phase (Fig. B). As above, for a critical comparison the results of several separate experiments were combined. Cells in individual experiments were divided into groups according to DNA content. For each group the topo IIα levels for all Ras-injected cells were compared to topo IIα levels for uninjected, neighboring cells. Results from all six experiments were combined to determine the mean ratio of the topo IIα level in injected cells to that in uninjected cells (± SE) for each DNA group (Fig. C). The topo IIα levels for injected and uninjected cells were remarkably similar, particularly during the first half of the cell cycle, although a slight reduction might have been induced by Ras during G2 phase. Similar results were obtained when Ras-injected cells were compared to cells injected with rat immunoglobulin as a control.
Ras stimulation of topo IIα in slowly cycling cells.
To confirm the above results, oncogenic Ras was injected into cells, which were then monitored for 25 h in time-lapse prior to fixation and analysis of topo IIα levels. As above, no overall stimulation of topo IIα levels by Ras could be observed (Fig. B). However, this analytical approach allowed the identification of cells with lengthened cell cycle phases. Cells which failed to divide in the 25 h of the time-lapse observation are represented as having an age of 25 h. The topo IIα levels in these cells were low as expected, except in the Ras-injected cells. In some of these slowly cycling, Ras-injected cells the topo IIα levels increased dramatically (Fig. B). Such cells would not have been apparent without time-lapse analysis. These cells are considered potentially interesting because they might serve as a culture model for tumor cells. Like most cells of a tumor, they retain proliferative capacity but cycle slowly. The fact that these cells responded to Ras injection by increasing topo IIα levels, therefore, is an observation which was carefully verified.
A second analysis of slowly cycling cells was performed by comparing ras-transformed and untransformed NIH 3T3 cells in asynchronous culture. Each cell type was cultured with [3H]thymidine for 20 h prior to staining for topo IIα. Cells which cycle slowly enough to avoid passing through any part of S phase during this time would remain unlabeled. In the ras-transformed culture these unlabeled cells contained a significantly higher topo IIα level than the unlabeled, untransformed cells (Fig. A). This result confirms that the oncogenic signaling resulted in topo IIα elevation in the slowly cycling cells. As above, the difference between transformed and untransformed cells was due to a few noncycling ras-transformed cells which expressed extremely high topo IIα levels.
As a final confirmation that oncogenic signaling can lead to high topo IIα levels in slowly cycling cells, an analysis was performed on NIH 3T3 cells transformed by a separate oncogene, src. This activated tyrosine kinase functions upstream of cellular Ras and was expected to stimulate many of the same targets as Ras. The transformed cells were labeled with BrdU for 24 h and then stained for BrdU and topo IIα. For each cell the BrdU fluorescence was plotted versus the topo IIα fluorescence. Slowly cycling cells in this analysis would fail to incorporate BrdU. As above, a proportion of these slowly cycling cells displayed high topo IIα levels (Fig. B). While only a fraction of these cells expressed high topo IIα, the highest topo IIα content in the culture was found in these non-BrdU-labeled, slowly cycling cells. These combined results indicate that, while the topo IIα levels of actively cycling cells appear to be held in check by cell size and DNA levels, when a cell pauses in its proliferation for a period of time, Ras activity is able to stimulate the production of high levels of topo IIα protein.
Drug toxicity and the cell cycle.
Once the effect of the cell cycle and oncogenic activity on topo IIα levels was determined, it was possible to determine the influence of each of these factors on drug toxicity. This comparison was simplified by the fact that neither the cell cycle nor oncogenic activity altered topo IIα levels, so that the effect of each could be determined without concern that secondary effects on topo IIα levels were involved. Drug toxicity studies were performed in asynchronous cultures with two consecutive time-lapse movies of the same cells separated by a brief (80-min) treatment with etoposide (2 to 10 μM). From the first movie (24 h) the cell cycle position of each individual cell at the time of drug treatment was determined, while from the second movie (50 to 75 h) the fate of each treated cell was analyzed (Fig. ). This allowed analysis of the effects of the cell cycle on drug toxicity without treatments required to induce cell cycle synchrony. Cells were poisoned with 5-azacytidine (100 to 500 μM) or tritiated thymidine (2 mCi/ml) (8
) as controls in separate analyses.
FIG. 6 Experimental procedure for cell cycle toxicity analysis. In order to determine the effects of the cell cycle on drug toxicity, NIH 3T3 cells were monitored in time-lapse for 25 h, the final 80 min of which was in the presence of the drug. The drug was (more ...)
Drug-treated cells displayed a variety of fates in the 2 to 3 days following treatment. Cells were considered normal if they divided at least twice during this time. They were considered to have been poisoned if they failed to divide, passed through an abortive mitosis, or underwent a rapid cell death. The typical appearance of rapid cell death and abortive mitosis is depicted in Fig. (see the figure legend for a web site leading to video clips of these events). For abnormal cells, the failure to divide was most common within the first 3 days following drug treatment. In prolonged analyses, however, those cells which failed to divide in the first 3 days generally exhibited either abortive mitosis or rapid cell death within 6 days following treatment (data not shown). Many cells divided only once in the second time-lapse analysis. These cells were carefully monitored throughout and were found to behave similarly to normal cells but are not included in any of the calculations discussed below.
FIG. 7 Rapid cell death and abortive mitosis. (Top) A small proportion of NIH 3T3 cells treated with etoposide experienced a sudden, dramatic cell death. A typical event is pictured, with the relative times indicated. Note that the most dramatic effects took (more ...)
Because the number of cells analyzed in any given experiment was limited, each toxicity experiment was repeated a number of times and the highly consistent data from each analysis were combined to yield a cumulative profile (Fig. ). Etoposide treatment of NIH 3T3 cells (7 μM) was most toxic when administered between 8 and 16 h following mitosis, when the cells were from mid-S phase to early G2 phase. Cells treated during G1 phase were much more likely to remain normal than cells treated during this sensitive phase. Interestingly, the toxicity of etoposide was reduced as cells progressed late into G2 phase (Fig. A). The validity of these results is emphasized by the fact that this experiment was performed independently at least six times with highly similar results. As a control cells were treated in the same way with another toxic agent, 5-azacytidine. In this case there was little cell cycle-specific killing, but the small effect seen was opposite to that obtained with etoposide, with the greatest survival of cells during S and G2 phases (Fig. C).
FIG. 8 Cell cycle and drug toxicity. (A) Six individual experiments were performed as described for Fig. . In these experiments NIH 3T3 cells were treated with etoposide (7 μM) to determine cell cycle-specific toxicity. The age and fate (more ...)
To more specifically relate the results with etoposide to specific cell cycle stages, and as another control, cells were poisoned with tritiated thymidine, which was found to be specific for S phase toxicity as expected. The experimental strategy was exactly as described above. The results with labeled thymidine were dramatic, with an almost quantitative killing of cells in S phase and little toxicity in other cell cycle stages (Fig. D). Not only does this result emphasize the validity of our cell cycle analytical approach, it clearly identifies the position of S phase in cells from 5 to 13 h old (Fig. D). It is therefore apparent that the maximal toxicity of etoposide is seen not in a given cell cycle stage but rather is localized from mid-S phase to early G2 phase (Fig. E).
The cell cycle toxicity experiments described above were next confirmed by treatment of cells synchronized by mitotic shakeoff. Mitotic NIH 3T3 cells were collected, replated, and treated with etoposide or 5-azacytidine for 80 min at various times after replating. The effects of the drug treatment were then determined by counting the cells after 3 days. To determine the timing of S phase entry following mitotic detachment, a set of parallel cultures was labeled with tritiated thymidine at various times following replating (Fig. ). While the delay between mitosis and S phase was much greater following mitotic detachment than in actively cycling cells, the cell cycle-related toxicity profiles were similar. As with the asynchronous cells, etoposide-induced toxicity in the shakeoff cells increased beginning in approximately mid-S phase until it reached a maximum late in the cell cycle. With 5-azacytidine, the toxicity during S phase might have been somewhat reduced compared to that during other cell cycle phases (Fig. ). From these studies it can be concluded that the cell cycle has an important effect on etoposide toxicity, with greatest sensitivity during late S and G2 phases.
FIG. 9 Drug toxicity after mitotic selection. At 90-min intervals mitotic NIH 3T3 cells were detached mechanically from asynchronous cultures and replated in several parallel plates. At the indicated times following replating each culture was treated with the (more ...) Oncogenic signaling and toxicity.
Experiments were last performed to determine the effect of oncogenic activity on etoposide toxicity. Experiments similar to those described above were repeated with NIH 3T3 cells transformed by the src
oncogene. As previously reported (5
), these cells were much more sensitive to etoposide toxicity than untransformed cells. Experiments were performed with 2 μM etoposide so that the proportion of cells killed would be approximately equal to that observed when untransformed cells were treated with 7 μM etoposide. While the sensitivity of src
-transformed cells increased, the overall cell cycle characteristics of killing were similar to those observed with untransformed cells, with the possible exception that the overall cell cycle effects of toxicity might have been reduced in the transformed cells (Fig. B). To extend this observation, NIH 3T3 cells were injected with oncogenic Ras protein in a defined area of the culture and the toxicity of etoposide (2 μM) was determined with the time-lapse analysis. In this case the overall proportions of normal cells for the injected and uninjected neighboring cells were determined. As a control, similar experiments were performed by microinjecting rat immunoglobulin as a negative control. In repeated experiments, a dramatically increased sensitivity of Ras-containing cells was observed. The proportion of poisoned cells increased over threefold following Ras injection (Fig. A). Unfortunately, the overall numbers of these cells were too small to make definite conclusions regarding the cell cycle profile of toxicity following Ras injection. Since we know that the overall levels of topo IIα do not increase following Ras injection, this clearly indicates that oncogenic activity sensitizes the cells to etoposide independently of alterations in topo IIα levels.
FIG. 10 Oncogenic transformation and etoposide toxicity. (A) The time-lapse analysis of toxicity (Fig. ) was applied to cells injected with oncogenic Ras (Leu61; 1 mg/ml). The proportion of the cells which divided normally or which were killed (more ...)
A final consideration relates to those few cells which failed to pass through mitosis prior to treatment in the above time-lapse toxicity studies (Fig. ). These cells were considered interesting because they responded to Ras injection by increasing topo IIα to high levels in some cases and because they are considered to be a potential culture model for cells of a tumor, most of which also cycle slowly. While the numbers of these cells were small, when the total number from all the numerous experiments described above were considered, it appeared that src transformation greatly increased the etoposide toxicity in these slowly cycling cells (Fig. B). This is interesting, since it is known that the topo IIα levels increase in only a small proportion of slowly cycling src-transformed cells, yet a high proportion of these cells displayed increased toxicity to etoposide treatment. This again emphasizes the role of oncogenic transformation in potentiating drug toxicity independently of topo IIα levels, although final conclusions from the slowly cycling cells await further work to identify increased numbers of such cells.